FimR and FimS: Biofilm Formation and Gene Expression inPorphyromonas gingivalis�
Alvin Lo,1 Christine Seers,2 Stuart Dashper,2 Catherine Butler,2 Glenn Walker,2 Katrina Walsh,2Deanne Catmull,2 Brigitte Hoffmann,2 Steven Cleal,2 Patricia Lissel,2
John Boyce,3 and Eric Reynolds2*Laboratory for Structural & Molecular Microbiology, Structural Biology Brussels—VUB, VIB Building E, 4th Floor, Pleinlaan 2,
1050 Brussels, Belgium1; Cooperative Research Centre for Oral Health Science, Melbourne Dental School and theBio21 Institute of Molecular Science and Biotechnology, The University of Melbourne, 720 Swanston Street,
Victoria 3010, Australia2; and Department of Microbiology, Monash University, Victoria, 3800, Australia3
Received 8 September 2009/Accepted 16 December 2009
Porphyromonas gingivalis is a late-colonizing bacterium of the subgingival dental plaque biofilm associatedwith periodontitis. Two P. gingivalis genes, fimR and fimS, are predicted to encode a two-component signaltransduction system comprising a response regulator (FimR) and a sensor histidine kinase (FimS). In thisstudy, we show that fimS and fimR, although contiguous on the genome, are not part of an operon. Weinactivated fimR and fimS in both the afimbriated strain W50 and the fimbriated strain ATCC 33277 anddemonstrated that both mutants formed significantly less biofilm than their respective wild-type strains.Quantitative reverse transcription–real-time PCR showed that expression of fimbriation genes was reduced inboth the fimS and fimR mutants of strain ATCC 33277. The mutations had no effect, in either strain, on the P.gingivalis growth rate or on the response to hydrogen peroxide or growth at pH 9, at 41°C, or at low heminavailability. Transcriptome analysis using DNA microarrays revealed that inactivation of fimS resulted in thedifferential expression of 10% of the P. gingivalis genome (>1.5-fold; P < 0.05). Notably genes encoding sevendifferent transcriptional regulators, including the fimR gene and three extracytoplasmic sigma factor genes,were differentially expressed in the fimS mutant.
Two-component signal transduction systems (TCSTS) areused by bacteria to control the expression of a range of genesin response to a variety of environmental and intracellularstimuli. These systems are found in almost all bacteria and areknown to regulate an array of physiological traits, includingosmoregulation (4), virulence (8), and quorum sensing (21).The crucial role of TCSTS in governing the signaling andregulatory pathways associated with biofilm development hasbeen well documented in many bacteria, including Escherichiacoli, Pseudomonas aeruginosa, and Streptococcus mutans (11,25, 39). Typically, each TCSTS functions via a phosphorylationcascade and consists of a membrane-bound or cytoplasmicsensor histidine kinase (SHK), which perceives a particularstimulus, and a cytoplasmic response regulator (RR), whichallows the cell to respond to the stimulus accordingly via reg-ulation of gene expression (49).
Porphyromonas gingivalis is a Gram-negative anaerobe thathas been strongly implicated as a major etiologic agent in theonset and progression of chronic periodontitis (47, 55), a dis-ease of the supporting tissues of the teeth. P. gingivalis is a latecolonizer of subgingival dental plaque, a complex and dynamicpolymicrobial biofilm (22, 31), and its ability to persist as partof a subgingival plaque biofilm is dependent on its adherenceto and colonization of the subgingival niche. P. gingivalis hasbeen shown to adhere to primary plaque-colonizing species,
particularly Streptococcus spp. such as Streptococcus gordonii(24, 29, 33, 48). Binding of P. gingivalis to S. gordonii has beenshown to result in the formation of a bispecies biofilm with P.gingivalis attached to S. gordonii bound to a salivary pellicle (7).P. gingivalis also adheres to later-colonizing Gram-negativebacteria, including Fusobacterium nucleatum (33, 43, 44) andTreponema denticola (14, 56). Furthermore, P. gingivalis canadhere to host tissues, including gingival epithelial cells (6, 23,40, 53). Therefore, to colonize and persist within a host P.gingivalis must sense the presence of a variety of surfaces andrespond via coordinated gene expression. Given the variationin surfaces to which P. gingivalis may attach, it is likely thatnumerous cell structures are required to mediate the interac-tions necessary for specific and stable adherence. Indeed, re-cent studies have shown that both the major fimbrillin FimAand the minor fimbrillin Mfa1 are required for full P. gingivalisbiofilm development by strain ATCC 33277 (26). In addition,the involvement of capsular polysaccharide and lipopolysac-charide O antigen has been implicated in P. gingivalis biofilmformation (10, 35). P. gingivalis W50 is afimbriated (53), so themechanism of biofilm formation for this strain is unclear.While stable attachment is clearly critical for the establishmentof P. gingivalis in the subgingival niche, continued survival atthis site requires appropriate bacterial responses to a range ofadverse conditions, including oxidative and nutrient stresses aswell as variations in temperature and pH. It is likely that manyof these responses are regulated by TCSTS.
Bioinformatics analysis of the P. gingivalis W83 genome se-quence (36) identified 6 putative TCSTS, one of which, GppX,is a predicted fusion of both SHK and RR proteins (16). A
* Corresponding author. Mailing address: Centre for Oral HealthScience, Melbourne Dental School, The University of Melbourne, 720Swanston Street, Victoria 3010, Australia. Phone: 61 3 9341 1547. Fax:61 3 9341 1596. E-mail: [email protected].
study comparing the transcriptomes of biofilm and planktonicP. gingivalis strain W50 cells identified two genes, pg1431 andpg1432, that were highly upregulated (4.2-fold and 5.2-fold,respectively) (27) during biofilm growth. The pg1431 gene ispredicted to encode a 227-amino-acid, 25.5-kDa putativeDNA-binding response regulator of the LuxR family, while thepg1432 gene is located upstream of, and in the same orienta-tion as, pg1431 and is predicted to encode a 621-amino-acid,70.1-kDa putative sensor histidine kinase. Genes homologousto pg1431 and pg1432 have been identified previously in P.gingivalis strain ATCC 33277 and were designated fimR andfimS, respectively (17); therefore, we designate the W50pg1431 gene as fimR and the W50 pg1432 gene as fimS. It hasbeen assumed that FimR (RR) and FimS (SHK) work inconcert as a two-component regulatory system. Inactivation ofeither fimS or fimR in strain ATCC 33277 resulted in loss offimbriation (17). Furthermore, FimR has been shown to di-rectly regulate the expression of a limited number of genes,including pg2130, which is associated with FimA fimbriation(37), and mfa1, which encodes the minor fimbrillin Mfa1 (54).The genes associated with the function of FimS have not beenexperimentally determined.
Here, we demonstrate that both FimR and FimS are in-volved in P. gingivalis biofilm formation, including the regula-tion of genes associated with fimbriation. Furthermore, DNAmicroarray analysis of a W50 fimS mutant suggests that thisSHK has a broad role in P. gingivalis gene regulation.
MATERIALS AND METHODS
Bacterial strains, growth conditions, and plasmids. The bacterial strains andplasmids used in this study are listed in Table 1. E. coli strain JM109 (Promega,Madison, WI) was grown in Luria-Bertani (LB) broth or on LB agar plates at37°C under aerobic conditions. Freeze-dried cultures of P. gingivalis strains W50and ATCC 33277 were obtained from the culture collection of The MelbourneDental School, The University of Melbourne. P. gingivalis strains were grown andmaintained as previously described (46). Growth media were supplemented with10 �g ml�1 erythromycin (Sigma) or 100 �g ml�1 of ampicillin (Sigma) whenappropriate.
P. gingivalis was grown in continuous culture for 30 days, in duplicate, using aBioflo 110 fermentor with a total volume of 400 ml (New Brunswick Scientific,Edison, NJ), as previously described (9). Planktonic cells were harvested fromthe fermentor by rapidly pumping them out, and the RNA was harvested usingan acidic hot phenol procedure (27). Culture purity was assessed regularly byGram staining and colony morphology.
DNA analysis and manipulation. Oligonucleotide primers used in this studyare listed in Table 2. Genomic DNA from P. gingivalis strains W50 and ATCC33277 and mutant strains were prepared using the DNeasy blood and tissue kit(Qiagen, Valencia, CA), and plasmid DNA from E. coli was extracted using theQiagen Miniprep kit (Qiagen). The Pfu DNA polymerase and restriction endo-nucleases (Promega) and Platinum Taq DNA polymerase High Fidelity (Invitro-gen Life Technologies, Carlsbad, CA) were used according to the manufacturer’sinstructions. Sequencing of DNA was performed by Applied Genetic Diagnos-tics, The University of Melbourne. Sequence alignments were done with theClustalW program (http://www.ebi.ac.uk/Tools/clustalw2/) (50). Where a PCRamplicon was sequenced, two separate PCRs were performed to ensure sequenceconsensus.
The fimR loci from strains W50 and ATCC 33277 were each amplified by PCRusing the oligonucleotide primer pair PG1432Seq-For and PG1430Seq-Rev,which annealed to the flanking genes. The W50 fimS locus was amplified usingPG1433Seq-For and PG1431Seq-Rev, while the strain ATCC 33277 fimS locuswas amplified using the oligonucleotide primer pair PG1431Seq-Rev andfimS_Seq-For. The resulting amplicons were purified using the QIAquick PCRpurification kit (Qiagen), and the nucleotide sequences were determined.
Construction of P. gingivalis fimR and fimS null mutants. To make the fimRmutagenesis cassette, a 353-bp DNA fragment containing the 5� region of fimR,with flanking AatII and BamHI restriction sites, was generated by PCR using P.gingivalis W50 DNA as template and the oligonucleotide primers PG1431-AatII-For and PG1431-BamHI-Rev. This amplicon was digested with AatII andBamHI and ligated into the AatII and BamHI sites, adjacent to the ermF genewithin pAL30 (9), to create pAL31. Similarly, a 264-bp DNA fragment contain-ing the 3� region of fimR, with flanking KpnI and SpeI restriction sites, wasamplified using the oligonucleotides PG1431-KpnI-For2 and PG1431-SpeI-Rev2and ligated into the KpnI and SpeI restriction sites in pAL31. The resultingplasmid, designated pAL31.1, had the ermF cassette flanked by fimR DNA.Plasmid pAL31.1 was linearized with ScaI and transformed into P. gingivalisstrains W50 and ATCC 33277 by electroporation as previously described (12).Transformants were selected after 7 days of anaerobic incubation on horse bloodagar plates containing 10 �g ml�1 of erythromycin. Gene disruptions wereconfirmed by PCR.
A similar strategy for fimS inactivation was followed, in which the 5� and 3�regions of fimS were amplified by PCR using the primer pairs PG1432-AatII-Forand PG1432-BamHI-Rev (5� region) and PG1432-KpnI-For2 and PG1432-SpeI-Rev2 (3� region). These fragments were sequentially ligated to pAL30, to finally
TABLE 1. Bacterial strains and plasmids used in this study
Bacterial strain or plasmid Relevant phenotypea, description, or selective marker Source or reference
Porphyromonas gingivalisW50 Wild type Laboratory collectionATCC 33277 Wild type Laboratory collectionECR220 P. gingivalis W50; fimR::ermF Emr This studyECR221 P. gingivalis ATCC 33277; fimR::ermF Emr This studyECR222 P. gingivalis W50; fimS::ermF Emr This studyECR223 P. gingivalis ATCC 33277; fimS::ermF Emr This study
PlasmidspVA2198 E. coli-Bacteroides shuttle vector carrying ermF-ermAM cassette; Spr 12pGEM-T Easy Cloning vector; Apr PromegapAL30 1,177-bp ermF cassette with incorporated BamHI and KpnI sites in pGEM-T Easy 9pAL31 353-bp upstream fragment of PG1431 ligated between AatII and BamHI sites of pAL30 This studypAL31.1 264-bp upstream fragment of PG1431 ligated between KpnII and SpeHI sites of pAL31 This studypAL32 831-bp upstream fragment of PG1432 ligated between AatII and BamHI sites of pAL30 This studypAL32.1 762-bp downstream fragment of PG1432 ligated between KpnI and SpeI sites of pAL32 This study
a Spr, spectinomycin resistant; Apr, ampicillin resistant; Emr, erythromycin resistant.
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produce pAL32.1. This plasmid was introduced into strains W50 and ATCC33277 by electroporation, and transformants were selected as described above.
RT and qRT-PCR. Total RNA for reverse transcription-PCR (RT-PCR) washarvested from P. gingivalis W50 chemostat-grown planktonic cells (27) and fromexponential-phase ATCC 33277 batch culture cells following the protocol ofDashper et al. (9). The total RNA (1 �g of each) was reverse transcribed tocDNA using the Superscript III first-strand synthesis Supermix kit (Invitrogen)with random hexamer oligonucleotide primers. The cDNA (10 ng) was used asthe template for PCR using BIOTAQ Red DNA polymerase (Bioline, Alexan-dria, Australia) with genomic DNA (10 ng) as a PCR-positive control. TotalRNA that had not been subjected to reverse transcription was used as a controlto show that the RT-PCR amplicons had not resulted from amplification ofcontaminating genomic DNA.
For quantitative real-time RT-PCR (qRT-PCR) analysis, total RNA was har-vested from at least three separate cultures of each P. gingivalis strain, ATCC33277, ECR221, and ECR223, during exponential-phase and stationary-phasegrowth in brain heart infusion (BHI) medium batch culture. RNA was harvestedfollowing the protocol of Dashper et al. (9) with cDNA synthesis and qRT-PCRsusing 100 ng of input RNA and the Superscript III Platinum 2-step qRT-PCRSYBR green kit reagents and protocol (Invitrogen, Van Allen Way, CA). TheqRT-PCRs were carried out using a Rotor-Gene 3000 instrument (Qiagen,Sydney, Australia). Melting curve analysis was performed in the temperaturerange from 50 to 99°C in 0.2°C increments. A no-RT control was used in eachrun. The mRNA abundance was determined by comparing the values obtainedfrom the qRT-PCR to a standard curve generated using ATCC 33277 genomeDNA. The expression of the housekeeping gene galE (27) was used for normal-ization between samples.
DNA microarray analysis. Total bacterial RNA, harvested from duplicateW50 and ECR222 chemostat-grown planktonic cultures, was reverse transcribedto cDNA and then labeled with Cy3 and Cy5 for use in DNA microarrayhybridizations with P. gingivalis oligonucleotide arrays as described previously
(27). Four DNA microarrays (kindly provided by the Pathogens FunctionalGenomics Resource Centre; http://pfgrc.jcvi.org) were used for each biologicalreplicate comparison, with a dye-swap design, making a total of eight slides usedin the analysis. Scanned images of the hybridized arrays were analyzed usingImagene 6.0 software (Biodiscovery, Los Angeles, CA) with local backgroundcorrection. Intensity-dependent Lowess normalization was applied usingGeneSight 4.1 (Biodiscovery), as described previously (27). Differentially ex-pressed genes were identified at 95% confidence intervals with a fold changethreshold value of 1.5. All DNA microarray work in this study was in compliancewith MIAME guidelines.
Mutant strain growth kinetics. Strains were initially grown overnight in ananaerobe chamber, and an aliquot of cells (1 � 109 CFU ml�1) was added to (i)fresh BHI medium for growth kinetics and temperature stress assays, (ii) BHImedium supplemented with 1 mM H2O2 (H2O2 stress), (iii) 0.1 �g ml�1 insteadof 5 �g ml�1 of hemin (hemin limitation stress), or (iv) BHI medium adjusted topH 5.0 or pH 9.0 (pH stress). The cells (260 �l) were transferred to triplicatewells of a microtiter plate (Falcon 353072; Becton Dickinson, North Rye, NSW,Australia), which was then sealed and inserted into a microtiter plate reader(Labsystems iEMS reader MF; Labsystems, Helsinki, Finland) and incubated at37°C. The growth of each strain was monitored by measurement of the opticaldensity at 620 nm (OD620). Growth curves were constructed using the mean andstandard deviation of three separate assays. Wells containing BHI medium onlywere used as blank controls.
Static biofilm formation assays. Static biofilm formation was assayed using theprotocol of O’Toole and Kolter (38) with slight modification. Briefly, an over-night culture was diluted with fresh BHI medium to obtain 5 � 107 CFU ml�1.The cells were aliquoted into the wells of a 96-well microtiter plate (260 �l perwell) and incubated anaerobically at 37°C for 24 h. The supernatant of theculture was aspirated, and then the well was washed twice with phosphate-buffered saline (150 mM NaCl, 3 mM KCl, 10 mM Na2HPO4, and 1.5 mMKH2PO4, pH 7.4). The biofilms were stained by incubation of each well with 100
TABLE 2. Oligonucleotides used in this study
Oligonucleotide Sequence (5�33�)a
DNA sequencing analysisPG1432Seq-For ..............................................................................................................................CGCGACTAACTATCCTGACAPG1430Seq-Rev .............................................................................................................................GGTTCGCTGACGAAACGTTTGTAGGCPG1433Seq-For ..............................................................................................................................TGAGTTCTTCGATCTGCTTGTCGGCTPG1431Seq-Rev .............................................................................................................................TGCCAATCCACTAATCCGCTfimS_Seq-For ..................................................................................................................................CACGGCTGTTCAGGTGGGCT
Construction of fimR mutantermF-BamHI-For ...........................................................................................................................CGCGGATCCCCGATAGCTTCCGCTATTermF-KpnI-Rev...............................................................................................................................GCCGGTACCTCCATCGCCAATTTGCCAPG1431-AatII-For..........................................................................................................................TGATTAGACGTCTACTCGTGGATGACCACGPG1431-BamHI-Rev......................................................................................................................AGAAATGGATCCCCGTGGTTACTGTGCGGAPG1431-KpnI-For2 ........................................................................................................................TACTGGGGTACCGCGTATCACGTTCCGCAGPG1431-SpeI-Rev2.........................................................................................................................CTATACTAGTTGCCAATCCACTAATCCGCT
Construction of fimS mutantPG1432-AatII-For..........................................................................................................................TTAATAGACGTCCGCGACTAACTATCCTGACPG1432-BamHI-Rev......................................................................................................................ATAGGATCCTACCCGGCATGGCATGCPG1432-KpnI-For2 ........................................................................................................................ATCCAAGGTACCGGCAGACTCTATTGCCGCPG1432-SpeI-Rev2.........................................................................................................................TCTTACTAGTATCCGATTCGAGGATATCGG
RT and qRT-PCR analysisLuxR-For ........................................................................................................................................CGCAGACCAATCGCATAAGLuxR-Rev........................................................................................................................................CAGAATAGCCATCGCACAGASenHis-For......................................................................................................................................CCATGCAGCAAGGAGATACASenHis-Rev .....................................................................................................................................TAGTGTCGAGGGCCATTTTC1431R2.............................................................................................................................................CGAGCAATATTCACTCCATTCfimAF...............................................................................................................................................AAGGTAATGCCACCATCAGCfimAR..............................................................................................................................................GCATTTTCGGCTGATTTGATmfa1F...............................................................................................................................................ACTTCTCCCGATTCATGGTGmfa1R..............................................................................................................................................GGATTCGGGTCAGGGTTATTpg0178F...........................................................................................................................................TATCGGGATCGTCCTCTTTGpg0178R ..........................................................................................................................................TAACCTCCCGAAACATCGAGgalEF ...............................................................................................................................................TCGGCGATGACTACGACACgalER...............................................................................................................................................CGCTCGCTTTCTCTTCATTC
a Restriction endonuclease target sites are underlined.
�l of 0.1% crystal violet (CV) for 5 min. The plate was then washed twice withdistilled water and destained with 95% ethanol (200 �l per well) for 5 min. Thesolubilized CV was transferred to a new microtiter plate, and the OD540 wasmeasured. Biofilm formation was qualitatively determined to be proportional tothe absorbance of the CV.
Microscopic analyses of static biofilms. Static biofilms were generated in a16-well chambered coverglass system (Grace Biolabs) as described previously (5)and stained using the BacLight bacterial viability assay kit (Invitrogen) accordingto the manufacturer’s instructions. The biofilms were examined using a ZeissLSM 510 Meta confocal laser scanning microscope (CLSM) with a C-Apochro-mat 63�/1.2 numerical aperture, water immersion objective lens with correctioncollar. SYTO 9 fluorescence (green, live cells) was detected by excitation at 488nm, and emission was collected with a 500- to 550-nm bandpass filter. Propidiumiodide (PI) fluorescence (red, dead cells) was detected by excitation at 488 nm,and emission was collected with a 560-nm long-pass filter. All images wereobtained over an area 142.9 � 142.9 �m in the x-y plane (parallel to the surface).z-stack images were obtained by taking serial optical slices in this plane over arange of distances at a resolution of 1,024 by 1,024 pixels. Three independentstatic biofilm experiments were performed for each strain, and at least 9 imagestacks were acquired for each experiment. These image stacks were quantita-tively analyzed using COMSTAT software (18) (The Math Works, Inc., Natick,MA) to determine the biomass, mean thickness, and roughness measurements ofthe biofilms formed by all strains.
Statistical analyses. Biofilm parameters obtained using confocal microscopyand transcript levels measured using qRT-PCR were statistically analyzed usinga one-way analysis of variance (ANOVA) with Scheffe’s post hoc multiple com-parison (32). For all statistical tests, � was set at 0.05. Levene’s test was per-formed to investigate homogeneity of variance. All statistical analyses wereperformed using Statistical Package for the Social Sciences, version 16.0.
Adherence of FITC-labeled P. gingivalis to KB cells. The binding of fluoresceinisothiocyanate (FITC)-labeled P. gingivalis to KB cells was carried out as de-scribed by Pathirana et al. (40). Bound bacteria were detected using an FC500flow cytometer (Beckman Coulter, Gladesville, NSW, Australia). KB cells wereidentified as a homogeneous population of large granular cells, and the bacteriaadhering to these cells were identified by fluorescence detected through a525-nm band-pass filter and compared with unlabeled controls.
Microarray data accession number. All new microarray data have been de-posited in the ArrayExpress databases under accession no. E-TABM-546.
RESULTS
Genetic and functional organization of fimR and fimS.Bioinformatic analysis of the predicted W83 strain FimR (36)using the Simple Modular Architecture Research Tool (SMART;http//smart.embl-heidelberg.de) (45) identified it as typical re-sponse regulator with a putative receiver domain between aminoacid residues 2 and 117 and a helix-turn-helix Lux regulon-type
DNA binding domain between amino acid residues 161 and218 (Fig. 1). SMART analysis of the W83 strain FimS identi-fied an N-terminal sequence typical of a long leader peptide(residues 1 to 40); however, this overlapped a second, shorter,predicted transmembrane domain (residues 21 to 40). With analternative methionine start codon at residue 19, this secondtransmembrane domain may also serve as the leader peptide.This suggests that FimS may be secreted across the innermembrane. Another transmembrane domain was predicted be-tween residues 375 and 397 followed by a histidine kinase Aphosphoacceptor domain (residues 412 to 479) and a histidinekinase-like ATPase domain (residues 524 to 619). Therefore,the predicted structure of FimS indicates that the sensor do-main is localized to the periplasm and linked to the cytoplas-mic kinase domain via transmembrane residues 375 to 397(Fig. 1). The predicted sensor domain has two tetratricopep-tide repeats (TPR; residues 143 to 176 and 183 to 216) and acoiled-coil region (residues 338 to 365), motifs that are in-volved in protein-protein interactions and protein stabilizationrespectively.
We determined the nucleotide sequence of the fimR andfimS loci from the P. gingivalis W50 (afimbriated) and ATCC33277 (fimbriated) strains. The W50 fimS locus was amplifiedby PCR using the oligonucleotide primers PG1433Seq-For andPG131Seq-Rev. However, these oligonucleotide primers didnot amplify a product from ATCC 33277. This suggested thatthe sequences upstream of the W50 fimS and the ATCC 33277fimS are divergent. To confirm this, we designed a forwardprimer oligonucleotide (fimS_Seq-For) based on the publishedATCC 33277 fimS locus sequence (17), repeated the PCR, andsequenced the amplicon. The result confirmed the data ofHayashi et al. (17) and showed that the nucleotide sequencesupstream of the fimS genes of the two strains are significantlydifferent (Fig. 2A). Indeed, the nucleotide sequence alignmentindicates that these genes may use different start codons andthe expressed proteins may have different N-terminal se-quences, although both can have possible leader peptides con-sistent with the PG1432 SMART residue 21-to-40 transmem-brane domain prediction. Importantly, the sequences of the
FIG. 1. Graphical representation of the functional domains of FimR and FimS identified by the Simple Modular Architecture Research Tool(SMART; http//smart.embl-heidelberg.de). In FimR, the symbols labeled “REC” and “HTH LUXR” denote the receiver domain (residues 2 to117) and the helix-turn-helix DNA binding Lux regulon domain (residues 161 to 218), respectively. In FimS, from left to right, the horizontal barsrepresent the putative signal peptide (residues 1 to 40), the low-complexity region (residues 263 to 274), and the coiled-coil region (residues 338to 365), respectively. The vertical bar represents a putative transmembrane segment (residues 375 to 397). A second putative transmembranesegment was predicted at residues 21 to 40 of FimS that overlaps the signal peptide prediction and is not shown in this figure. Two Pfam-predictedtetratricopeptide repeats, TPR_1 and TPR_2 (residues 143 to 176 and 186 to 216) are also shown. The amino acids at positions 412 to 479 and524 to 619 compose the putative histidine kinase A phosphoacceptor domain (HisKA) and the histidine kinase-like ATPase domain (HATPase_c),respectively.
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promoters and putative regulatory elements upstream of thesefimS genes must differ, which has implications for the tran-scriptional regulation of these loci in the different strains.
In addition to these differences, a single base transition wasdetected within the region encoding the histidine kinase do-mains of FimS, changing the coded amino acid from isoleucineto lysine in the W50 FimS (Fig. 2B). This change is not locatedwithin the predicted phosphoacceptor or the histidine kinase-like ATPase domains and is unlikely to affect the histidinekinase function. More significant, however, was the presence ofan additional adenosine base in the W50 fimS at 1,843 bp fromthe predicted PG1432 ATG start codon that was not present inthe ATCC 33277 fimS (Fig. 2B). This altered reading framewould result in a shorter FimS product in strain W50. Thesequenced W50 fimR and fimS genes are identical to those ofgenes pg1431 and pg1432 of strain W83.
fimR and fimS do not constitute an operon. Typically, genesencoding the response regulator and the sensor histidine ki-nase of two-component signal transduction systems are contig-uous and transcribed together in an operon. It has been pre-sumed but not verified that fimS and fimR, which are separatedby only 12 nucleotides in the ATCC 33277 genome and 65
nucleotides in the W50 genome (Fig. 2B) are cotranscribed. Toexplore this possibility, the transcription of fimR and fimS ineach strain was analyzed by RT-PCR. Using oligonucleotidesspecific for fimR (LuxR-For and LuxR-Rev) or fimS (SenHis-For and SenHis-Rev) gave RT-PCR products of 113 bp and131 bp, respectively, as expected (Fig. 3). However, PCR witha primer pair designed to span the intergenic region betweenfimR and fimS (SenHis-For and PG1431R2) yielded no ampli-cons (Fig. 3, lanes 14 and 17), whereas PCR using the sameoligonucleotides with genomic DNA as the template gave anamplicon of 954 bp, as expected (Fig. 3, lanes 16 and 19).Taken together, these results show that fimR and fimS are notcotranscribed. Furthermore, these data indicate that theremust be a promoter specific for fimR expression between fimSand fimR. We identified a sequence motif, TAGGTTTG, thatis similar to the highly conserved �7 sequence motif, TAnnTTTG, found as part of the consensus P. gingivalis and otherBacteriodetes promoter sequences (2, 19, 30), and there is alsoan alternative fimR start codon 18 bp downstream of this se-quence (Fig. 2B).
The role(s) of FimR and FimS in P. gingivalis biofilm for-mation. To explore the significance of the FimR and FimS in
FIG. 2. Partial alignment of the DNA and amino acid translations of P. gingivalis strain W50 and ATCC 33277 fimS and fimR genes and theencoded products. (A) Alignment of the 5� regions (uppercase) of the sequenced W50 fimS gene and the published ATCC 33277 fimS gene(GenBank accession no. AB025360). The predicted translation start codons (ATG) of W50 fimS and ATCC 33277 fimS are in boldface andunderlined, and the upstream untranslated regions are in lowercase. Nucleotide numbering is from the A of the putative ATG translation startcodons, which is number 1. Identical nucleotides are marked with an asterisk. Alternative putative start codons are shaded gray. (B) Alignmentof the 3� regions of the sequenced P. gingivalis W50 and ATCC 33277 fimS genes and the 5� region of the fimR genes. The intergenic nucleotidesare in lowercase. The encoded products presented below the nucleotide sequences show the K-to-I substitution in the fimS products (in boldfaceand underlined) and the fimS frameshift introduced by presence of nine A bases (W50 fimS and our laboratory strain of ATCC 33277) comparedwith eight A bases reported in AB025360. The W50 fimS DNA sequence was identical to that of P. gingivalis strain W83 sequence (www.tigr.org).Highlighted in gray are the database-predicted methionine start codons and the downstream alternative start codons of fimR. Underlined is aputative �7 promoter motif. Identical nucleotides are marked with an asterisk.
P. gingivalis biofilm formation, fimR and fimS of strains ATCC33277 and W50 were each disrupted by insertion of an ermFcassette. The W50 fimR mutant was designated ECR220, theATCC 33277 fimR mutant was designated ECR221, the W50fimS mutant was designated ECR222, and the ATCC 33277fimS mutant was designated ECR223.
The ability of each of the fimR and fimS mutants to formbiofilms was initially examined using a rapid 24-h, 96-well mi-crotiter plate static biofilm assay. ATCC 33277 formed mea-surable CV-stained biofilms in this assay; however, P. gingivalisstrain W50 did not, even though this strain is known to producebiofilms in continuous culture (1, 27). In an attempt to induceW50 to form biofilms in this system, the microtiter wells werecoated with filter-sterilized human saliva (5 to 10 �g of pro-tein) or fibronectin from human plasma (5 to 10 �g), but thesestrategies were unsuccessful. The wells were also coated with5 � 107 formalin-killed S. gordonii or F. nucleatum cells priorto seeding with P. gingivalis and overnight incubation. How-ever, incubation with P. gingivalis W50 caused the S. gordoniiand F. nucleatum cells to release from the wells. As we wereunable to find conditions amenable to P. gingivalis W50 biofilmformation in this system, we confined the rapid biofilm studiesto ATCC 33277 and the ATCC 33277 mutants ECR221 andECR223.
The fimR and fimS mutants ECR221 and ECR223 bothformed significantly less biofilm (P � 0.001, t test) than theATCC 33277 parent strain (Fig. 4), with ECR221 and ECR223having, respectively, 81% and 60% less CV-staining biofilmbiomass than ATCC 33277. Furthermore, the biofilm biomassformed by ECR221 was significantly smaller than that of thefimS mutant ECR223 (P � 0.001). To investigate the possibil-ity that the reduced biofilm formation by ECR221 andECR223 could be attributed to reduced growth rate, thegrowth kinetics of the mutants and the wild-type strains werecompared. The mean generation times for the mutants(ECR223, 5.0 � 0.4 h; ECR221, 4.4 � 0.2 h) were not signif-
icantly different from that of the wild type (ATCC 33277, 4.6 �0.5 h). Furthermore, at 24 h, the time point at which biofilmformation was assessed, ECR221 and ECR223 achieved opti-cal densities (0.98 � 0.03 and 0.90 � 0.01, respectively) thatwere comparable to that of the ATCC 33277 wild type (0.96 �0.02). Therefore, the reduced biofilm formation by strainsECR221 and ECR223 could not be attributed to reducedgrowth rates.
Analysis of the biofilms formed by P. gingivalis ATCC 33277,ECR221, and ECR223 by confocal laser scanning microscopy.Confocal laser scanning microscopy was used to analyze thearchitecture of the biofilms formed by ATCC 33277, ECR221,and ECR223. Strains were grown in a 16-well culture chambercoverglass system and stained using BacLight live-dead stain.Both ECR221 and ECR223 formed biofilms that were visiblymore sparse than the biofilm formed by the parent ATCC33277, with the ECR221 biofilm being composed of smaller,more dispersed aggregates in comparison to ECR223 (Fig. 5).Quantitative analysis using COMSTAT (Table 3) showed thatthe calculated biomasses of ECR221 and ECR223 biofilmswere significantly reduced (P � 0.01) to 7% and 25%, respec-tively, of that produced by the wild-type strain ATCC 33277.The mean thickness of the biofilm produced by each mutant(0.3 � 0.12 �m for ECR221 and 1.33 � 0.30 �m for ECR223)was also significantly reduced relative to ATCC 33277, whichwas 4.52 � 1.31 �m thick (P � 0.01). However, both mutantsdisplayed similar maximum thickness to that of the wild type(Table 3), suggesting that microcolony tower formation wasnot limited, but rather there was a less effective establishmentof intercolony contacts in these biofilms. This supposition wassupported by the roughness coefficients (R) of 1.8 and 1.2 forECR221 and ECR223 biofilms, respectively, indicating the for-mation of rough and heterogenous biofilms that are oftenassociated with many pillars and towers of cells separated byareas devoid of cells (18). The roughness coefficients of thebiofilms produced by each of the mutants were significantly
FIG. 3. RT-PCR analysis of fimR and fimS transcription in W50and ATCC 33277. RT-PCR was performed using primer pairs de-signed to amplify within the fimS and fimR coding regions or to spanthe intergenic region of fimR and fimS. Total RNAs from P. gingivalisW50 chemostat-grown planktonic cells and ATCC 33277 batch-grownplanktonic cells were reverse transcribed, and the cDNAs were used asPCR templates. Lane 1, Gene Ruler 50-bp DNA ladder (Fermentas,Glen Burnie, MD); lanes 2 to 7, primer pair specific for fimR; lanes 8to 13, primer pair for fimS; lanes 14 to 19, primer pair designed to spanthe intergenic region of fimR and fimS; lane 20, broad-range DNAladder (Marligen Biosciences, Ijamsville, MD). Lanes 2, 8, and 14,W50 cDNA PCR templates; lanes 5, 11, and 17, ATCC 33277 cDNAPCR templates. The following controls were used: lanes 3, 9, and 15,W50 total RNA, no RT; lanes 6, 12, and 18, ATCC 33277 total RNA,no RT; lanes 4, 10, and 16, W50 genomic DNA; and lanes 7, 13, and19, ATCC 33277 genomic DNA.
FIG. 4. Biofilm formation by P. gingivalis ATCC 33277 and thefimR (ECR221) and fimS (ECR223) mutant strains in 96-well micro-titer plates. Biofilms were stained with crystal violet and destainedusing ethanol. The absorbance of the crystal violet in the ethanolfraction was measured at 540 nm and was proportional to the P.gingivalis biofilm biomass formed. The data shown are representativeof three independent assays in which triplicate biofilm samples foreach strain were measured. The results are expressed as the mean, anderror bars indicate 1 standard deviation. Asterisks indicate a statisti-cally significant difference (P � 0.001; t test) in biofilm between P.gingivalis ATCC 33277 and the mutants (ECR221 and ECR223) as wellas between each of the mutants.
VOL. 192, 2010 FimR AND FimS IN P. GINGIVALIS BIOFILM 1337
different (P � 0.01), indicating that the mutants producedbiofilms with dissimilar morphologies. In contrast, the wild-type biofilm had a very low R (0.08), indicating that this strainforms a homogenous and uniform biofilm (Table 3).
The expression of genes important for P. gingivalis fimbri-ation in the fimS and fimR mutants. It has been shown that
production of the fimbrillins Mfa1 and FimA is essential forbiofilm formation by P. gingivalis ATCC 33277 (26). Further-more, it has been shown that FimR binds directly to the mfa1promoter region (54) and to the promoter of the fimbriation-associated gene fimX that is upstream of fimA (37) (annotatedas pg2130 in the W83 genome [36] and pgn_0178 in the recentlyreleased ATCC 33277 genome sequence [34]). We used qRT-PCR to quantify mfa1, pgn_0178, and fimA transcripts in thefimS and fimR mutants and ATCC 33277, normalizing thetranscript levels relative to that of the control gene galE (27).
In comparison to wild-type cells, the expression of mfa1,fimA, and pgn_0178 in each of the mutants was decreased, butto different extents for each gene. The level of fimA andpgn_0178 expression was significantly reduced, by at least 28-fold, in the fimS and fimR mutants (P � 0.05), from levels upto 35-fold greater than galE expression in the wild type to levelsbelow galE expression in the mutants (Fig. 6). Notably, in thewild-type strain ATCC 33277 the level of FimA-encoding tran-script was 9-fold higher than that of the pgn_0178 transcript,indicating that transcription of more fimA mRNA is initiatedfrom the fimA proximal promoter than from the promoterproximal to pgn_0178. In the fimS and fimR mutants, althoughthe abundances of the pgn_0178 and fimA mRNAs were bothvery low, this differential transcript abundance between fimAand pgn_0178 continued, with the ratio being 19-fold in thefimR mutant and 13-fold in the fimS mutant. However, therewas no statistically significant difference between the level ofeither fimA or pgn_0178 expression in each of the mutants.
The decrease in expression of the minor fimbrillin-encodinggene mfa1 in the fimS and fimR mutants was less profound,although statistically significant. Interestingly, the expressionof mfa1 was found to be influenced by growth phase. In ATCC33277, the level of mfa1 expression in exponential-phase cells(OD650 of 0.5 to 0.6) was significantly higher (1.9-fold; P �0.01) than that measured in stationary-phase cells (OD650 of1.4). Similarly, in the fimR mutant there was a 1.6-fold differ-
FIG. 5. CLSM analysis of the biofilms formed by P. gingivalis ATCC 33277 (A) and the fimR (ECR221) (B) and fimS (ECR223) (C) mutants.Biofilms were grown in a 16-well culture chamber coverglass system and stained with SYTO 9 and PI. The z-stack of each was acquired by CLSMwith a C-Apochromat 63�/1.2 water immersion objective lens with corrrection collar. The dimensions of the region displayed are 512 �m by 512�m. The data presented here are representative of three independent experiments.
TABLE 3. Quantitative analysis of the P. gingivalis ATCC 33277,ECR221, and ECR223 biofilm architectures
Parameter andsignificance comparisona
Result for strain:
ATCC 33277 ECR221 ECR223
Biomass (�m3/�m2)b 4.10 � 1.22 0.29 � 0.14 1.10 � 0.50P value for ATCC
33277 vs mutant0.003 0.009
P value for ECR221 vsECR223
NSc
Roughness coefficientb 0.08 � 0.03 1.80 � 0.08 1.23 � 0.23P value for ATCC
33277 vs mutant�0.001 �0.001
P value for ECR221 vsECR223
0.009
Avg thickness (�m)b 4.52 � 1.31 0.30 � 0.12 1.33 � 0.30P value for ATCC
33277 vs mutant0.002 0.007
P value for ECR221 vsECR223
NS
Maximum thickness(�m)b
5.47 � 1.40 3.87 � 0.23 4.67 � 1.40
P value for ATCC33277 vs mutant
NS NS
P value for ECR221 vsECR223
NS
a All P values were determined using one-way ANOVA with Scheffé’s post hocmultiple comparison tests.
b Values are expressed as means � standard deviations.c NS, not significant (P � 0.05).
ence in mfa1 expression (P � 0.05) between exponential- andstationary-phase cells (Fig. 6). In contrast, in the fimS mutantthere was no significant change in the level of mfa1 transcriptbetween the exponential and stationary phases. In the expo-nential growth phase, the fimS mutant expressed decreasedlevels of mfa1 relative to wild-type cells (2.7-fold; P � 0.001),a difference that decreased to 1.5-fold in the stationary phase.In the fimR mutant, the decrease in mfa1 expression was notstatistically significant. There was no significant difference be-tween the levels of mfa1 expression in any of the strains duringthe stationary phase of growth.
Overall, the data clearly show that FimR and FimS eachhave a function in the control of expression of genes importantfor P. gingivalis fimbriation and that each affects the expressionof these genes to differing extents. Reduction in fimbriationdue to decreased fimbriation-associated gene expression mayin part explain the reduced capacity of the ATCC 33277 FimRand FimS mutants to form biofilms in vitro.
Binding of P. gingivalis to KB cells. The P. gingivalis W50wild-type strain and fimR and fimS mutants were grown tomid-exponential phase, harvested, labeled with FITC, and in-cubated with KB cell monolayers. P. gingivalis strain ATCC33277 was not used for the KB cell binding assays due to thepropensity of the strain to self-aggregate (40). The percentageof KB cells with bound P. gingivalis was determined using flowcytometry. There was no difference in the numbers of KB cellswith bound P. gingivalis W50 wild-type or fimR/fimS cells, andmean fluorescence intensities were also equivalent (data notshown), indicating that equal numbers of mutant and wild-typeP. gingivalis cells were bound per KB cell (data not shown).Therefore, this suggests that the binding of P. gingivalis W50 toKB cells was not dependent on expression of fimR or fimS.
Stress assays. The involvement of FimR and FimS in the P.gingivalis response to various physiological stresses such asH2O2, elevated temperature, hemin limitation, and altered pHwas investigated. The fimR and fimS mutants, in the presenceof 1 mM H2O2, showed no difference in growth kinetics fromtheir respective wild-type strains (ATCC 3327 and W50), norwas any difference observed under temperature stress (42°C),hemin limitation (0.1 mg ml�1), or growth at altered pH (datanot shown). Thus, under the test conditions used, neitherFimR nor FimS had any significant role in the responses of P.gingivalis W50 or ATCC 33277 to H2O2, hemin limitation,elevated temperature, or altered pH.
The effect of the disruption fimS on the global gene expres-sion of P. gingivalis. To gain further insight into the role ofFimS in P. gingivalis gene expression, we used DNA microar-rays to compare the transcriptome of the wild-type strain, W50,with that of the fimS mutant ECR222. As the W50 fimS mutantwould not grow as a biofilm, we used wild-type and fimS mu-tant (ECR222) cells grown in planktonic culture for the micro-array analysis.
Relative to strain W50, the disruption of fimS resulted in thealtered expression (�1.5-fold up- or downregulated; P � 0.05)of 199 genes in ECR222 cells, of which 110 genes showedincreased expression and 89 genes exhibited decreased expres-sion relative to the parent strain (see Tables S1 and S2 in thesupplemental material). This represents 10% of the P. gingiva-lis genome. The level of fimR expression was reduced by 2.3-fold in the fimS mutant. The signal intensity of the hybridizedfimS probe was equal to the array background signal intensity,confirming that there was no expression of fimS in ECR222.These data indicate that fimR expression is influenced by, butnot completely codependent on, fimS transcription and sup-
FIG. 6. Expression of fimA, pgn_0178, and mfa1 in ATCC 33277 (wild type [wt]) fimS and fimR mutants as determined by qRT-PCR. Thehousekeeping gene galE was used to normalize values between replicates. Total RNA was harvested from at least three separate cultures of eachstrain, and qRT-PCR was performed using three technical replicates for each RNA sample. All expression is reported as the fold difference relativeto the measured amount of galE. *, significant differences in fimA and pgn_0178 expression in the fimS and fimR mutants relative to wild-type cells(P � 0.05); **, significant difference in mfa1 expression in wild-type cells between the exponential and stationary phases (P � 0.01); #, significantdifference in mfa1 expression in the fimR mutant between the exponential and stationary phases (P � 0.05); ##, significant difference in mfa1expression between the wild type and the fimS mutant during exponential-phase growth (P � 0.001).
VOL. 192, 2010 FimR AND FimS IN P. GINGIVALIS BIOFILM 1339
port the RT-PCR analysis that these genes are not cotrans-cribed.
In agreement with the qRT-PCR findings for the ATCC33277 fimS mutant ECR223, there was reduced expression ofthe fimX (pgn_0178) homologue pg2130 (13-fold) and fimAorthologue (pg2132) (6.5-fold). It is interesting to note that thefimA orthologue is expressed in W50 and the protein monomerdetected by proteomic analysis (9), but the fimbriae are notassembled. Also downregulated were other fimbriation-associ-ated genes (pg2131 and 2133-pg2136) (Table 4). In the genomesequence of strain W83 (36), mfa1 is interrupted by insertionelement ISPg4. We used PCR and DNA sequencing andshowed that mfa1 is also interrupted by ISPg4 in strain W50(data not shown). The transcript that would be the truncatedmfa1 mRNA (pg0176) was downregulated in the array 3-fold,in agreement with the reduced mfa1 expression measured byqRT-PCR in the ATCC 33277 fimS mutant. The disruption offimS also resulted in the upregulation of most of the genes ofa 6-gene cluster, PG0508 to PG0513, that also has reducedexpression in P. gingivalis mature biofilms (27), suggesting thatFimS negatively affects the expression of these genes in bothbiofilm and planktonic cells through an as-yet-unidentified re-pressor. Also potentially derepressed was PG0718, encoding aconserved hypothetical protein (16-fold upregulation), andPG0862, encoding a putative restriction endonuclease (12-foldincrease in expression).
When the differentially regulated genes were grouped intoTIGR (www.tigr.org) role categories (Fig. 7), more than half ofthe differentially expressed genes observed in ECR222 (55genes upregulated and 48 genes downregulated) encoded hy-pothetical proteins, conserved hypothetical proteins, or pro-teins with similarity to proteins with uncharacterized functions.
Of the other functional categories, the expression of genes en-coding proteins predicted to be involved in the binding and trans-port of substrates was most affected in the fimS mutant (23 geneswere up- and downregulated, respectively, in ECR222) (Table 4).Genes involved in amino acid biosynthesis and fatty acid andphospholipid metabolism were apparently unaffected by the dis-ruption of fimS.
TABLE 4. Selected genes that were differentially expressed in the fimS mutant ECR222
TIGR openreading frame Gene name Annotation Cellular role Fold change in
FIG. 7. Number of genes with altered (by 1.5-fold or more; P �0.05) expression in the P. gingivalis W50 fimS mutant ECR222 groupedby TIGR functional category. A, amino acid biosynthesis; B, biosyn-thesis of cofactors, prosthetic groups, and carriers; C, cell envelope; D,cellular processes; E, central intermediary metabolism; F, DNA me-tabolism; G, disrupted reading frame; H, energy metabolism; I, fattyacid and phospholipid metabolism; J, mobile and extrachromosomalelement functions; K, protein fate; L, protein synthesis; M, purines,pyrimidines, nucleosides, and nucleotides; N, regulatory functions; O,signal transduction; P, transcription; Q, transport and binding proteins;R, unknown function; and S, hypothetical or conserved hypotheticalproteins.
During the establishment of a biofilm and during biofilmgrowth, cells must remain responsive to environmental changesand be able to maintain an appropriate balance between theplanktonic and biofilm growth phases, a process that requiresthe interplay of various cell factors. As a late colonizer ofsubgingival plaque, P. gingivalis persistence is dependent on itsability to stably attach to a variety of surfaces, including otherbacteria and the subgingival plaque extracellular matrix, aswell as possibly epithelial cells of the subgingival crevice. Giventhat each of these surfaces will have distinct properties, it islikely that P. gingivalis must use specific extracellular protein(s)to facilitate these diverse interactions. Thus, P. gingivalis wouldneed to regulate the production of an array of extracellularproteins to adapt to the various surfaces it encounters. Thismay be reflected in the activation of fimS, fimR, and genesassociated with the biogenesis of a cell envelope componentsuch as the FimA and Mfa1-type fimbriae when P. gingivalis isgrown as biofilm (27).
Two-component signal transduction systems have been iden-tified and characterized in various bacterial species (49). Rolesfor some of these TCSTS in biofilm development, as well asother phenotypic traits, have been demonstrated in bothGram-negative and Gram-positive bacteria, and the regulatorysystems often display remarkable complexity. For example,biofilm formation by E. coli involves production of curli (11).Curli production is influenced by the CpxA-CpxR sensor ki-nase-response regulator system (11) but also by CsgD, EnvZ/OmpR, Rcs, and H-NS (20). Furthermore, the regulator CsgDinfluences cell aggregation and cellulose production and affectsthe expression of at least 24 genes, including other regula-tors (3).
Species that have a restricted ecological niche, such as in-tracellular parasites, have few sensor proteins encoded withintheir genomes, whereas in contrast, species such as Mesorhizo-bium loti, P. aeruginosa, and Vibrio cholerae, which occupymore variable habitats have more than 100 sensor proteins(13). The detection of only 6 putative sensor histidine kinasesin the P. gingivalis W83 genome suggests that it may be pro-grammed to sense few environmental factors. The independenttranscription of fimS and fimR may provide added flexibility tothe response regulation cascades in P. gingivalis with FimSand/or FimR able to interact with alternative sensor histidinekinases and response regulators in a manner that would not bepossible if they were transcriptionally linked. The concept thatFimS interacts with response regulators other than FimR issupported by the microarray data that showed change in theexpression of a large number of genes following the disruptionof fimS in ECR222 (10% of the genome). In comparison,transcriptomic analysis of a fimR mutant revealed a relativelysmall regulon with only 7 genes identified as differentially ex-pressed (37). Our microarray data revealed that the expressionof four predicted transcriptional regulators (PG0173, PG0543,PG1044, and PG2125) was downregulated in ECR222, whiletwo other regulators (PG0928 and PG1181) were upregulated.In addition, three putative RNA polymerase extracytoplasmicsigma-70 factors (PG0214, PG0985, and PG1827) were up-regulated in ECR222. These data strongly suggest that FimS ispart of a complex cascade of regulatory effectors. Cross-talk
and cross-regulation phenomena, albeit a new concept with P.gingivalis, have been documented in other species, including E.coli and Pseudomonas spp. (42, 52). Given the limited numberof TCSTSs in the P. gingivalis genome, the broad specificity ofFimS may increase the adaptability of the response of theorganism to environmental change.
Expression of fimS also had a negative influence on theexpression of genes with very diverse cellular functions. Thus,we propose a role for FimS as the major physiological “switch”as part of P. gingivalis biofilm development, with a major effectbeing reduction in fimbriation. The importance of FimS in P.gingivalis biofilm formation coupled with the fact that fimS ishighly upregulated during P. gingivalis mature biofilm growth(27) suggests that fimS may be activated during both of thesedistinct phases of P. gingivalis biofilm development.
The disruption of either fimS or fimR of ATCC 33277 re-sulted in significant impairment (but not abolition) of biofilmformation by P. gingivalis in 96-well microtiter plate and glassculture chamber systems. Furthermore, a W50 fimS mutantalso formed only sparse, poorly attached biofilm in a fermentorvessel. CLSM analysis of the biofilms formed by the ATCC33277 fimR and fimS mutants showed that they produced bio-films that were distinct from the wild-type strain and also fromeach other, with rough and heterogeneous biofilms suggestingan apparent reduction of microcolony contacts. In vitro analysisof ATCC 33277 fimA and mfa1 mutants has shown FimA to beinvolved in adhesion of P. gingivalis to saliva-coated glass sur-faces (26) and cultured KB epithelial cells (51), while mfa1expression mediates cell aggregation (26). Neither the fimAnor mfa1 mutants formed a confluent biofilm (26). In view ofthis, reduced expression of fimA and mfa1 in the ATCC 33277fimS and fimR mutants, with resulting lowered FimA and Mfa1production, would explain the biofilm architecture we observedusing CLSM and would also explain the lowered biofilm pro-duction by these mutants. However, this is not sufficient toexplain the impairment of biofilm formation by the afimbriatedstrain W50, indicating that there are factors other than fimbri-ation that are involved in the initiation of P. gingivalis biofilmformation. As the distinct physiological steps of P. gingivalisbiofilm development have yet to be fully elucidated, it is prob-able that in concert with fimbriae, the products of the extensivenumber of hypothetical genes shown to have altered expres-sion in ECR222 have a role in P. gingivalis biofilm formation.
P. gingivalis W50 has previously been shown to bind to KBcells (40) and fibroblasts (41), with the RgpA-Kgp cysteineproteinase-adhesin complexes produced by P. gingivalis havingsome role in this adherence. Using isogenic mutants, it hasbeen shown that production of the Kgp proteinase had themost significant role in this effect (40, 41). We could measureno difference in the adherence of strain W50 and the ECR222fimS mutant to KB cells, and the microarray data indicatedthat there was no change in the expression of kgp or rgpA,which encode the components of the RgpA-Kgp complexes.Measurement of Arg- and Lys-specific whole-cell proteinaseactivity also showed no difference between the strains (data notshown). Together, these data indicate that strain W50 KBepithelial cell adhesion is mediated by factors other than thoseassociated with FimS function and that FimS does not functionin the regulation of the expression of rgpA or kgp. Interestingly,other than a slight upregulation of hagA (2.1-fold), none of the
VOL. 192, 2010 FimR AND FimS IN P. GINGIVALIS BIOFILM 1341
genes that encode the numerous surface-associated CTD fam-ily proteins to which RgpA and Kgp belong (46), many ofwhich are adhesins, were differentially expressed in ECR222,indicating that FimS does not function to regulate expressionof genes encoding these surface proteins.
The qRT-PCR data showed that, during exponential growth,mfa1 expression was decreased more in the fimS mutant thanin the fimR mutant, suggesting that there may be at least oneother activator of mfa1 expression which is dependent on phos-phorylation by FimS. Possible candidate transcription factorsrevealed by the microarray data are PG0543, MntR (PG1044),PG2125, and PG0173, as all of the genes encoding these pro-teins were downregulated in ECR222 (Table 4).
The presence of an N-terminal sequence typical of a signalpeptide followed by two transmembrane helices indicates thatthe sensor domain of FimS with the TPR is likely to be local-ized to the periplasm of the cell (Fig. 1). TPR domains arefound to be widely distributed from prokaryotes to eukaryotesand are known to act as molecular scaffolds in mediating spe-cific protein-protein interaction (15), while coiled-coil struc-tures are known to facilitate and stabilize protein-protein in-teractions (28). Given this and the result that FimS was highlyinduced in a P. gingivalis W50 biofilm (27), we propose thatFimS may sense a signal that is important in the developmentof P. gingivalis biofilm. We observed no altered ability of theFimS mutants to respond to a range of stresses that P. gingivaliswould experience during biofilm growth in the subgingivalniche, including H2O2 (which may result from neutrophil at-tack), temperature changes (as occurs during inflammation),hemin limitation, and altered pH.
Concluding remarks. To our knowledge, this is the firststudy to directly show the important involvement of both FimSand FimR in P. gingivalis biofilm development. We have dem-onstrated that fimR and fimS are not part of an operon andalso that fimS and fimR mutants form altered biofilm pheno-types compared to both wild-type strains and each other. Incontrast to the situation observed in an fimR mutant, wherealtered regulation of only a limited number of genes was ob-served, the disruption of fimS resulted in the altered expressionof a large number of genes encoding products with very diversecellular functions. We hypothesize that FimS may be importantin monitoring the specific environmental signal that is requiredfor or associated with P. gingivalis fimbriation and growth as abiofilm and forms part of a complex regulatory network regu-lating biofilm formation and development.
ACKNOWLEDGMENTS
This work was funded by National Health and Medical Researchgrant no. 509326 and Australian Dental Research Foundation grant44/2006.
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